A novel mechanism by which histone deacetylase inhibitors facilitate topoisomerase IIα degradation in hepatocellular carcinoma cells

Abstract

Histone deacetylase (HDAC) inhibitors exhibit a unique ability to degrade topoisomerase (topo)IIα in hepatocellular carcinoma (HCC) cells, which contrasts with the effect of topoII-targeted drugs on topoIIβ degradation. This selective degradation might foster novel strategies for HCC treatment in light of the correlation of topoIIα overexpression with the aggressive tumor phenotype and chemoresistance. Here, we report a novel pathway by which HDAC inhibitors mediate topoIIα proteolysis in HCC cells. Our data indicate that HDAC inhibitors transcriptionally activated casein kinase (CK)2α expression through increased association of acetylated histone H3 with the CK2α gene promoter. In turn, CK2 facilitated the binding of topoIIα to COP9 signalosome subunit (Csn)5 via topoIIα phosphorylation. Furthermore, we identified Fbw7, a Csn5-interacting F-box protein, as the E3 ligase that targeted topoIIα for degradation. Moreover, siRNA-mediated knockdown of CK2α, Csn5, or Fbw7 reversed HDAC inhibitor-induced topoIIα degradation. Mutational analysis indicates that the ¹³⁶¹SPKLSNKE¹³⁶⁸ motif plays a crucial role in regulating topoIIα protein stability. This motif contains the consensus recognition sites for CK2 (SXXE), glycogen synthase kinase (GSK)3β (SXXXS), and Fbw7 (SPXXS). This study also reports the novel finding that topoIIα may be a target of GSK3β phosphorylation. Evidence suggests that CK2 serves as a priming kinase, through phosphorylation at Ser1365, for GSK3β-mediated phosphorylation at Ser1361. This double phosphorylation facilitated the recruitment of Fbw7 to the phospho-degron ¹³⁶¹pSPKLpS¹³⁶⁵ of topoIIα, leading to its ubiquitin-dependent degradation.

Conclusion

This study shows a novel pathway by which HDAC inhibitors facilitate the selective degradation of topoIIα, which underlies the complexity of the functional role of HDAC in regulating tumorigenesis and aggressive phenotype in HCC cells.

Hepatocellular carcinoma (HCC) is a leading cause of cancer death worldwide. The clinical management of HCC is complicated by typically late-stage disease at presentation and prevalent underlying liver dysfunction that can render patients ineligible for potentially curative surgical therapies, which are generally suitable for only 20%-30% of HCC patients (1). Although regional therapies, such as transarterial embolization and percutaneous treatments, are used in patients with nonresectable disease, their success is curtailed by recurrence as locally advanced or metastatic disease (2). For these patients, systemic therapies are indicated but have been largely unsuccessful, in part, due to cellular resistance to conventional cytotoxic agents (3, 4). Thus, a clear need exists to develop effective, life-prolonging therapeutic strategies for the large number of HCC patients with advanced disease (5).

Previously, we demonstrated that the novel phenylbutyrate-derived histone deacetylase (HDAC) inhibitor AR42 (formerly OSU-HDAC42) exhibited high in vivo potency in suppressing HCC tumor growth, which was attributable to its ability to target both histone acetylation-dependent and –independent pathways (6). In addition to HDAC inhibition, AR42 also blocked the phosphorylation/expression level of a series of apoptotic regulators, including Akt, Bcl-xL, survivin, cIAP1, and cIAP2. Here, we show that AR42 facilitates the proteasomal degradation of topoisomerase (topo)IIα without disturbing topoIIβ expression in HCC cells, which was also noted with MS-275, a class I HDAC inhibitor, and, to a lesser extent, vorinostat (suberoylanilide hydroxamic acid).

The unique ability of HDAC inhibitors to degrade topoIIα contrasts with the selective effect of topoII-targeted drugs on topoIIβ degradation (7,8), and may foster novel strategies for HCC treatment considering the correlation of topoIIα overexpression with the aggressive tumor phenotype and chemoresistance (9,10). Moreover, topoIIβ may underlie many of the side effects associated with topoII-targeted drugs, such as doxorubicin-induced cardiotoxicity (11) and etoposide-induced secondary malignancies (12). From a mechanistic perspective, HDAC inhibitors provide a useful tool to elucidate the pathways governing topoIIα degradation, which represents the focus of this study.

Experimental Procedures

Cell line, culture and reagents

PLC5 and HepG2 cells were obtained from the American Type Culture Collection (Manassas, VA), and Huh7 cells were from the Health Science Research Resources Bank (Osaka, Japan). These HCC cells were cultured in Dulbecco’s modified Eagle’s medium (Invitrogen, Carlsbad, CA) supplemented with 10% fetal bovine serum (Invitrogen). All cells were cultured at 37°C in a humidified incubator containing 5% CO₂. The HDAC inhibitors vorinostat, MS-275, and AR42 (OSU-HDAC42) (6,13,14) were synthesized in our laboratory with purities exceeding 99%. MG132, wortmannin, PD98059, SB202190, SB216763, and DMAT were purchased from Sigma-Aldrich (St. Louis, MO). Bay11-7082 and GF-109203X were from Calbiochem (San Diego, CA). Antibodies against various proteins were from the following sources: topoIIα, BD Transduction (San Diego, CA); topoIIβ, casein kinase (CK)2α, Ets-1, HDAC1, and HDAC6, Santa Cruz (Santa Cruz, CA); Fbw7, Bmi1 and Skp2, Invitrogen; Fbx4, Rockland (Gilbertsville, PA); Fbx7, ProteinTech (Chicago, IL); Flag, Sigma-Aldrich; β-actin, MP Biomedicals (Irvine, CA); COP9 signalosome subunit (Csn)5, GeneTex (Irvine, CA); p-Ser/Thr, Abcam (Cambridge, MA); acetyl-histone H3, Millipore (Billerica, MA). Goat anti-rabbit and rabbit anti-mouse IgG-horseradish peroxidase conjugates were from Jackson Laboratories (West Grove, PA).

Transient transfection and immunoblotting

PLC5 cells were transfected with Lipofectamine 2000 (Life Technologies, Gaithersburg, MD) according to the manufacturer’s protocol. Plasmids and RNA interference were obtained from the following sources: short-hairpin (sh)RNA constructs against HDAC1, HDAC2, HDAC6, and CK2α, and plasmids encoding CK2α and Csn5, Origene (Rockville, MD); small interfering (si)RNAs against Csn5, HDAC4, and HDAC5, Invitrogen; Fbw7 shRNA; Addgene. Immunoblotting was performed as previously described (14).

Co-immunoprecipitation analysis

Cells were treated with AR42 for 48 h and lysed by buffer B (5 mM HEPES, 1.5 mM MgCl₂, 0.2 mM EDTA, 0.5 mM DTT, 26% glycerol (v/v), 300 mM NaCl, pH 7.9) on ice for 1 h. After centrifugation at 13,000xg for 20 min, one-tenth volume of supernatant was stored at 4°C for use as input, and the remainder was incubated with protein A/G-Sepharose beads for 1 h to eliminate nonspecific binding. The mixture was centrifuged at 1,000xg for 5 min, and the supernatants were incubated with anti-topoIIα antibodies and protein A/G Sepharose overnight. The immunocomplexes were resolved by SDS-PAGE and proteins were detected with indicated antibodies.

Chromatin immunoprecipitation (ChIP) assay

PLC5 cells were treated with AR42 for 36 h, and fixed in 1% formaldehyde for 15 min to immobilize histone to DNA. Cross-linking was stopped with 125 mM glycine for 5 min. ChIP was performed as previously described (6) using antibodies against acetyl-histone H3 or Ets-1 with non-specific rabbit IgG as negative control. Primers spanning the proximal promoter regions of CK2α were used for amplification by reverse-transcription polymerase chain reaction (RT-PCR): 5′-GGGGATTCCTTCCATTTTGC-3′/5′-ATGGAGGAGGAGACACACGG-3′.

RT-PCR

Total RNA was isolated from drug-treated cells with Trizol reagent (Invitrogen) and chloroform extraction. Aliquots of 2 μg of total RNA were reverse-transcribed to cDNA with the iScript cDNA Synthesis Kit (Bio-Rad Laboratories, Hercules, CA) according to the manufacturer’s instructions. PCR products were resolved by agarose (1.2%) gel electrophoresis and visualized by ethidium bromide staining. The sequences of primers used were as follows: topoIIα, 5′-CGTCAGAACATGGACCCAGA-3′/5′-AGCAGATTCAGCACCAAGCA-3′; CK2α, 5′-TGAGGATAGCCAAGGTTCT-3′/5′-CAGCAGCAATCACTGGTGA-3′; GAPDH, 5′-AGGGGTCTACATGGCAACTG-3′/5′-CGACCACTTTGTCAAGCTCA-3′; HDAC1, 5′-CCACATCAGTCCTTCCAATA-3′/5′-TTCTCCTCCTTGGTTTTCTC-3′; HDAC6, 5′-CAACTGAGACCGTGGAGAG-3′/5′-CCTGTGCGGAGACTGTAGC-3′.

Plasmid construction and site-directed mutagenesis

Plasmids encoding various topoIIα mutations were generated from Flag-TopoIIα (GeneCopoeia, Rockville, MD) by site-directed mutagenesis using the QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA). Primers used to generate topoIIα mutations were as follows: S1361A, 5′-TGCTAGTCCACCTAAGACCAAAACTGCCCCAAAACTTAG-3′/5′-CTAAGTTTTGGGGCAGTTTTGGTCTTAGGTGGACTAGCA-3′; S1365A, 5′-GACCAAAACTTCCCCAAAACTTGCTAACAAAGAACTGAAACCACAG-3′/5′-CTGTGGTTTCAGTTCTTTGTTAGCAAGTTTTGGGGAAGTTTTGGTC-3′; E1368A, 5′-CCCCAAAACTTAGTAACAAAGCACTGAAACCACAGAAAAGTGT-3′/5′-ACACTTTTCTGTGGTTTCAGTGCTTTGTTACTAAGTTT-TGGGG-3′; S1393A, 5′-GGGCAGTGTACCACTGTCTTCAGCCCCTCCTGCTAC-3′/5′-GTAGCAGGAGGGGCTGAAGACAGTGGTACACTGCCC-3′; T1397A, 5′-CTTCAAGCCCTCCTGCTGCACATTTCCCAGATGAA-3′/5′-TTCATCTGGGAAATGTGCAGCAGGAGGGCTTGAAG-3′.

In vivo mechanistic validation

Female athymic nude mice (5-6 weeks of age) were obtained from Harlan Laboratories (Indianapolis, IN). All experimental procedures were done according to protocols approved by The OSU Institutional Laboratory Animal Care and Use Committee. Each mouse was injected subcutaneously with 1×10⁶ PLC5 cells in 0.1 mL serum-free medium containing 50% Matrigel. Mice with established tumors (mean starting tumor volume, 223 ± 75 mm³) were randomized to two groups (n = 5) that received the following treatments daily by gavage (10 μL/g body weight) for 3 or 6 days: (a) methylcellulose/Tween 80 vehicle, and (b) AR42 at 25 mg/kg. At the study endpoint, tumors were snap-frozen and stored at −80°C for subsequent co-immunoprecipitation analysis.

Results

Differential suppression of topoIIα expression by HDAC inhibitors

Pursuant to our finding that AR42 exhibits high in vivo efficacy against PLC5 tumor growth (6), we examined the effects of AR42 on various biomarkers pertinent to the aggressive phenotype of HCC, among which the concentration- and time-dependent suppression of topoIIα expression was noteworthy (Fig. 1A). As AR42 inhibited topoIIα expression at concentrations well below its IC₅₀ of 0.72 μM in inhibiting cell viability (6), this downregulation was not consequent to drug-induced cell death. This topoIIα repression was also noted with MS-275 and, to a lesser extent, vorinostat, however, at an-order-of-magnitude higher concentrations. This drug-induced suppression was topoIIα-selective since these HDAC inhibitors did not cause changes in topoIIβ expression. The suppressive effect of these HDAC inhibitors on topoIIα expression was also demonstrated in Huh7 and HepG2 cells (Fig. 1B).

Fig. 1.

Western blot analysis of the differential inhibition of topoIIα expression by HDAC inhibitors in HCC cells. (A) Upper panel, concentration-dependent effects of AR42, MS-275 and vorinostat on the expression of topoIIα versus topoIIβ after 48 h. Lower panel, time-dependent effect of 0.5 μM AR42 and 5 μM MS-275 on topoIIα versus topoIIβ expression in PLC-5 cells. (B) Concentration-dependent effects of AR42, MS-275, and vorinostat on topoIIα expression after 48 h in Huh7 and HepG2 cells. (C) Concentration-dependent, biphasic effect of sodium butyrate on topoIIα expression, with no appreciable effect on topoIIβ expression, in PLC-5 cells. The values in percentage, the average of two independent experiments, denote the relative intensities of protein bands of drug-treated samples to that of the DMSO-treated control after normalization to β-actin.

Published reports of the effects of other HDAC inhibitors on topoIIα expression indicate a cell type- and/or context-specificity. For example, treatment of D54 glioblastoma cells with trichostatin A or vorinostat had no effect on topoIIα expression (15). While sodium butyrate was reported to sensitize leukemia cells to etoposide by increasing topoIIα gene expression (16), treatment of MCF-7 cells with valproic acid led to transcriptional repression of topoIIα (17). To clarify this issue, we assessed the concentration-dependent effect of sodium butyrate on topoIIα expression in PLC5 cells. Our data show that treatment with a range of concentrations of sodium butyrate revealed a biphasic effect on topoIIα expression levels, i.e., upregulation at low concentrations (≤ 0.25 mM) and downregulation at higher concentrations (≥ 0.5 mM), without disturbing topoIIβ expression (Fig. 1C). These concentrations are consistent with those of sodium butyrate (0.4 mM) and valproic acid (2 mM) that upregulated and downregulated topoIIα expression, respectively, in the aforementioned studies. This dichotomous effect may typify the complex mode of action of short-chain fatty acids in regulating topoIIα expression relative to other HDAC inhibitors examined.

HDAC inhibitors promote topoIIα degradation

The finding that MS-275 was able to suppress topoIIα expression suggests the involvement of class I HDACs in the drug response. Thus, we assessed the effect of shRNA or siRNA-mediated knockdown of class I (HDAC1 and 2) vis-à-vis class II isozymes (HDAC4 - 6) on topoIIα mRNA and protein expression in PLC5 cells. Silencing of HDAC1 caused a sharp decrease in the topoIIα protein level, while the mRNA expression was not altered (Fig. 2A). However, the knockdown of other isozymes had no effect on the mRNA or protein expression of topoIIα. Evidence indicates that this topoIIα downregulation was attributable to proteasomal degradation. First, exposure of PLC5 cells to AR42 or MS-275 did not casue appreciable changes in topoIIα mRNA levels as determined by RT-PCR (Fig. 2B). Second, the proteasome inhibitor MG132 protected cells against the suppressive effect of AR42, MS-275, and vorinostat on topoIIα expression (Fig. 2C). Third, in the presence of cycloheximide, AR42 promoted the elimination of topoIIα relative to the DMSO control (Fig. 2D). Together, these data suggest a pivotal role of HDAC1 in the regulation of topoIIα protein stability.

Fig. 2.

Evidence that HDAC inhibitor-induced downregulation of topoIIα expression involves inhibition of HDAC1 and is attributable to proteasomal degradation in PLC5 cells. (A) sh/siRNA-mediated knockdown of HDAC1, but not HDAC2, 4, 5, and 6, decreased the protein level of topoIIα without affecting mRNA expression. (B) Assessment of the concentration- and time-dependent effects of AR42 and MS-275 on topoIIα mRNA expression levels as determined by RT-PCR. Neither drug affected topoIIα mRNA expression. (C) The proteasome inhibitor MG132 protected cells from HDAC inhibitor-induced topoIIα ablation. Cells were treated with HDAC inhibitors at the indicated concentrations for 24 h, followed by cotreatment with 5 μM MG132 for an additional 24 h. (D) Effect of AR42 on topoIIα protein stability. Cells were exposed to DMSO or 0.5 μM AR42 for 12 h followed by 100 μg/ml cycloheximide (CHX) for 6 or 12 h.

CK2 is involved in ubiquitin-dependent degradation of topoIIα

It is well documented that ubiquitin-dependent protein degradation is preceded by phosphorylation (18). As shown in Fig. 3A, concentration-dependent topoIIα repression by AR42 was accompanied by parallel increases in p-Ser/Thr phosphorylation and ubiquitination. However, no appreciable acetylation of topoIIα was noted in response to AR42 treatment, suggesting that topoIIα stability is not influenced by HDAC-regulated acetylation. Thus, to shed light onto the mechanism by which HDAC inhibitors facilitated topoIIα proteolysis, we first investigated the identity of the kinase involved in AR42-mediated topoIIα repression by examining the abilities of a panel of kinase inhibitors to block this cellular response. As CK2 (19,20), protein kinase Cδ (21), and extracellular signal-regulated protein kinase (22) have been reported to target topoIIα, we assessed the effects of their respective inhibitors, DMAT, GF-109203X, and PD98059, on AR42-induced topoIIα repression. Also, inhibitors of phosphoinositide 3-kinase (wortmannin), IκB kinase (Bay11-7082), and p38 MAP kinase (SB202190) were used as controls. Among them, DMAT exhibited a unique ability to block AR42-facilitated topoIIα repression, while the other inhibitors showed no appreciable protective effect (Fig. 3B). This finding suggests a mechanistic link between CK2, a tetrameric kinase comprised of two catalytic subunits (α and α’) and two identical regulatory subunits (β) (23), and HDAC inhibitor-mediated topoIIα proteolysis.

Fig. 3.

Potential role CK2 in AR42-induced topoIIα degradation in PLC5 cells. (A) Co-immunoprecipitation analysis of the concentration-dependent effect of AR42 on topoIIα acetylation (Ac-Lys), Ser/Thr phosphorylation (p-Ser/Thr) and ubiquitination (Ub). (B) Effects of inhibitors of different kinases on AR42 (0.5 μM)-induced topoIIα downregulation after 48 h.

CK2 forms a stable, catalytically active complex with topoIIα (20), and has been implicated in the modulation of topoIIα trafficking (24). Here, we obtained three lines of evidence to corroborate the role CK2 in promoting HDAC inhibitor-induced topoIIα degradation.

First, AR42 and MS-275 treatment led to concentration-dependent increases in CK2α protein and mRNA expression in PLC5 cells (Fig. 4A), suggesting the transcriptional activation of CK2α expression by HDAC inhibitors. ChIP analysis revealed that AR42 treatment caused a concentration-dependent increase in the association of CK2α promoter DNA with acetylated histone H3 (Fig. 4B), which in turn was associated with the enhanced recruitment of the transcription factor Ets-1, a key regulatory element of the CK2α gene (25), to the promoter, without altering the expression level of Ets-1 (Fig. 4C). Moreover, shRNA-mediated HDAC1 knockdown led to increased CK2α expression like that observed with topoIIα repression (Fig. 4D). Together, these findings provide direct evidence of the involvement of HDAC inhibition in the observed increase in CK2α expression. Second, overexpression of CK2α mimicked the suppressive effect of HDAC inhibitors on topoIIα expression without disturbing topoIIβ (Fig. 4E). Third, shRNA-mediated CK2α knockdown protected PLC5 cells from AR42- and MS-275-mediated inhibition of topoIIα expression (Fig. 4F).

Fig. 4.

Evidence that CK2α is involved in HDAC inhibitor-mediated topoIIα degradation. (A) Concentration-dependent effects of AR42 and MS-275 on the protein (upper panel) and mRNA (lower panel) expression levels of CK2α after 48 h. (B) Increased association of acetylated histone H3 (Ac-H3) with the CK2α gene promoter in AR42-treated cells as determined by ChIP analysis. (C) Upper panel, increased recruitment of Ets-1 to the CK2α gene promoter in response to AR42 treatment as determined by ChIP analysis. Lower panel, AR42 does not affect the global expression of Ets-1. (D) Effect of shRNA-mediated knockdown of HDAC1 on CK2α expression. (E) Dose-dependent effects of ectopically expressed CK2α on the expression of topoIIα and topoIIβ after 48 h. (F) Protective effects of shRNA-mediated silencing of CK2α against AR42 (AR)- and MS-275 (MS)-mediated topoIIα degradation. Cells were transfected with CK2α shRNA, incubated for 24 h, and treated with 0.5 μM AR42 or 5 μM MS-275 for 48 h.

Role of Csn5 in HDAC inhibitor-mediated topoIIα degradation

Csn5 [a.k.a., Jun-activation domain-binding protein-1 (Jab1)], a component of the COP9 signalsome complex, plays a critical role in the degradation of a number of signaling proteins (26). We hypothesized that Csn5 plays an intermediary role between increased CK2α expression and topoIIα degradation based on the following published data: (1) Csn5 facilitates topoIIα degradation in response to glucose starvation by interacting with topoIIα’s glucose-regulated destruction domain (27). (2) Csn5-mediated degradation of its target proteins can be prevented by the pharmacological inhibition of CK2, a Csn complex-associated kinase (28). These data, together with our findings, prompted us to investigate the involvement of Csn5 in the HDAC inhibitor-induced topoIIα degradation.

As shown in Fig. 5A, treatment of PLC5 cells with AR42 had no effect on Csn5 expression (input), but led to a concentration-dependent increase in the association of topoIIα with CK2α and Csn5 (right panel), which is noteworthy in that physical interaction with Csn5 is reported to be a prerequisite for the degradation of its target proteins (27). This increase in the amount of CK2α associated with the Csn5-topoIIα complex paralleled the increase in total cellular CK2α levels in AR42-treated cells. Moreover, the ectopic expression of Csn5 dose-dependently mimicked the suppressive effect of HDAC inhibitors on topoIIα expression (Fig. 5B), while siRNA-mediated knockdown of Csn5 protected against the drug-induced downregulation of topoIIα in AR42- and MS-275-treated PLC5 cells (Fig. 5C). These results are consistent with the putative role of Csn5 in HDAC inhibitor-mediated topoIIα degradation.

Fig. 5.

Evidence that Csn5 plays a crucial role in HDAC inhibitor-induced topoIIα degradation in PLC5 cells. (A) Co-immunoprecipitation analysis reveals the concentration-dependent, enhancing effect of AR42 on the association of CK2α and Csn5 with topoIIα. (B) Dose-dependent suppressive effect of the ectopic expression of Csn5 on topoIIα protein levels after 48 h of incubation. (C) siRNA-mediated silencing of Csn5 expression protects cells against AR42- or MS-275-induced topoIIα degradation. Cells were transiently transfected with scrambled (Scr) or Csn5-specific siRNA, and then treated with 0.5 μM AR42 or 5 μM MS-275 for 48 h.

Fbw7 acts as an E3 ligase that targets topoIIα for Csn5-induced degradation

The Csn complex facilitates the proteasomal degradation of target proteins by functioning as a docking platform for recruitment of the target’s specific kinase and E3 ligase (29). Consequently, we sought to identify the E3 ligase that targets topoIIα in the Csn5 complex. Csn5 is known to maintain the stability of a number of the F-box proteins of the Skp1–Cul1–F-box-protein family, including Skp2, Fbw7, Fbx4, and Fbx7, as the silencing of Csn5 led to the downregulation of these F-box proteins (30). Thus, using these Csn5-interacting F-box proteins as candidates for the topoIIα-targeted E3 ligase, we assessed the concentration-dependent effects of AR42 on the binding of these F-box proteins to topoIIα. The E3 ligase Bmi1 was also assessed in light of a recent report that Bmi1 controlled topoIIα degradation in response to glucose starvation (31).

PLC5 cells exhibited robust expression of Skp2, Fbw7, and Bmi1, but had low abundance of Fbx4 and Fbx7 (Fig. 6A, input). Co-immunoprecipitation revealed a concentration-dependent increase in the binding of Fbw7 to topoIIα by AR42 (right panel). This AR42-induced association was highly selective since the other F-box proteins were undetectable or present in extremely low levels, relative to Fbw7, in the complex formation with topoIIα. The functional role of Fbw7 as the topoIIα-targeted E3 ligase was further supported by the protective effect of shRNA-mediated knockdown of Fbw7 on AR42- and MS-275-mediated topoIIα ablation (Fig. 6B).

Fig. 6.

Evidence that Fbw7 acts as the E3-ligase mediating the degradation of topoIIα in HDAC inhibitor-treated PLC5 cells. (A) Co-immunoprecipitation analysis of the concentration-dependent effects of AR42 on the binding of different F-box proteins, including Skp2, Fbw7, Fbx4, Fbx7, and Bmi1, to topoIIα. (B) shRNA-mediated knockdown of Fbw7 protects cells from AR42- and MS-275-induced topoIIα degradation. Cells were transfected with the pRetrosuper (pRS) control vector or Fbw7 shRNA, and then exposed to 0.5 μM AR42 (AR) or 5 μM MS-275 (MS) for 48 h. (C) Pharmacological inhibition of CK2 by DMAT, as evidenced by lack of topoIIα phosphorylation, blocks the formation of the topoIIα-Fbw7-Csn5-CK2α complex. Left two panels: co-immunoprecipitation analysis of the concentration-dependent effects of AR42 on the association of topoIIα with Fbw7, Csn5, and CK2α. Right two panels: co-immunoprecipitation analysis of the concentration-dependent effects of AR42 in the presence of 5 μM DMAT on the association of topoIIα with Fbw7, Csn5, and CK2α. Cells were treated with AR42 alone or in combination with 5 μM DMAT for 48 h. (D) The presence of two putative Fbw7 recognition motifs (boxed) in the C-terminal domain of topoIIα. All mapped CK2 recognition sites are underlined, and the positions of Ser1361, Ser1365, Glu1368, Ser1393, and Thr1397 are indicated by numbers on top of these residues.

Above, we showed that, in addition to Csn5, CK2α also associated with topoIIα in response to AR42 (Fig. 5A). Thus, we hypothesized that phosphorylation of topoIIα by CK2 facilitated the association of topoIIα with the Csn5-Fbw7 complex in AR42-treated cells. Results in support of this hypothesis are shown in Fig. 6C, where the CK2 inhibitor DMAT abrogated the interaction of topoIIα with Csn5 and Fbw7. Exposure of PLC5 cells to AR42 induced a concentration-dependent increase in topoIIα phosphorylation, accompanied by parallel increases in its association with Csn5 and Fbw7, culminating in topoIIα proteolysis (left panel). However, pharmacological inhibition of CK2 by DMAT prevented increases above basal levels of AR42-induced topoIIα phosphorylation and its consequent association with Csn5 and Fbw7, thereby protecting topoIIα from drug-induced degradation (Fig. 6C, right panel).

Glycogen synthase kinase (GSK)3β-dependent binding of topoIIα to Fbw7 through a recognition motif at the C-terminus

Fbw7 recognizes the Cdc4 phosphodegron (CPD) motif of (S/T)PXX(S/T) (X, any amino acid) in many of its target proteins, including cyclin E, Myc, Jun, SV40 large T antigen, and the sterol regulatory element binding protein (32). Within this CPD motif, phosphorylation at the Thr residue by GSK3β in conjunction with that at the Ser residue by a priming kinase is required for binding. Analysis of the topoIIα sequence revealed two plausible Fbw7 recognition motifs, ¹³⁶¹SPKLS¹³⁶⁵ and ¹³⁹³SPPAT¹³⁹⁷ in the C-terminal domain (Fig. 6D, boxed). It is especially noteworthy that the former motif encompasses a well characterized GSK3β phosphorylation motif (SXXXS) and overlaps with a putative CK2 recognition site ¹³⁶⁵SNKE¹³⁶⁸ [consensus sequence, (S/T)XX(D/E); all mapped CK2 sites are underlined] (33), suggesting that CK2 might be the priming kinase for GSK3β-mediated phosphorylation of topoIIα.

The involvement of GSK3β in AR42-mediated topoIIα degradation was corroborated by several lines of evidence. First, pharmacological inhibition of GSK3β by SB-216763 protected cells against the suppressive effect of AR42 on topoIIα expression (Fig. 7A). Second, co-immunoprecipitation indicates that AR42 led to a concentration-dependent increase in the association of topoIIα with GSK3β (Fig. 7B). Third, ectopic GSK3β expression mimicked dose-dependently the effects of AR42 on the levels of topoIIα expression (Fig. 7C, left panel) and phosphorylation (right panel), and its association with Fbw7 (right panel).

Fig. 7.

Evidence that glycogen synthase kinase (GSK)3β is involved in AR42-mediated topoIIα degradation and identification of the Fbw7 recognition motif in topoIIα in PLC5 cells. (A) Protective effect of the GSK3β inhibitor SB-216763 on AR42-induced topoIIα downregulation. Cells were treated with 10 μM SB-216763 and 0.5 μM AR42 for 48 h. (B) Co-immunoprecipitation analysis reveals concentration-dependent increases in the binding of GSK3β to topoIIα after 48 h treatment with AR42. (C) Ectopic GSK3β expression dose-dependently increases topoIIα phosphorylation and its association with Fbw7. Cells were transfected with increasing doses of plasmid expressing hemagglutinin (HA)-tagged GSK3β, and after 48 h of incubation, cell lysates were immunoblotted. (D) Mutational analysis of the topoIIα residues Ser-1361, Ser-1365, Glu-1368, Ser-1393, and Thr-1397 to evaluate their roles in regulating CK2α-mediated topoIIα degradation and topoIIα binding to Fbw7. Cells were transfected with wild-type (WT) or mutant forms of Flag-tagged topoIIα (Flag-TopoIIα with or without co-transfection with CK2α plasmid, followed by co-immunoprecipitation analysis. (E) Co-immunoprecipitation analysis of the effect of DMAT (5 μM) and SB-216763 (10 μM) on AR42-induced association of topoIIα with CK2α and GSK3β Cells were treated with 0.5 μM AR42 plus 10 μM SB-216763 or 5 μM DMAT for 48 h.

The involvement of the ¹³⁶¹SPKLSNKE¹³⁶⁸ motif in regulating topoIIα protein stability through interactions with Fbw7, GSK3β and CK2 was supported by mutational analyses. Flag-tagged topoIIα mutants were created by replacing the Ser1361, Ser1365, Glu1368, Ser1393, or Thr1397 residue with Ala via site-directed mutagenesis, and then expressed in PLC5 cells in the presence or absence of ectopically expressed CK2α. Ectopic CK2α expression was used to mimic HDAC inhibitor-induced CK2α upregulation and consequent topoIIα degradation because treatment with AR42 and other HDAC inhibitors induced the expression of the transfected Flag-topoIIα (data not shown), presumably through the epigenetic activation of transcription. Of these five mutants, only S1361A, S1365A, and E1368A abrogated the suppressive effect of CK2α overexpression on topoIIα expression (Fig. 7D, input). Co-immunoprecipitation analysis indicates that this reversal of drug action was attributable to the inability of the S1361A, S1365A, and E1368A mutants to bind Fbw7 (Fig. 7D, lower panel). In contrast, S1393A and T1397 did not confer protection against CK2α-induced degradation or binding to Fbw7, indicating that the ¹³⁹³SPPAT¹³⁹⁷ motif did not play a role in mediating topoIIα degradation in the presence of ectopically expressed CK2α.

The premise that CK2 might be the priming kinase for GSK3β-mediated phosphorylation of topoIIα was supported by co-immunoprecipitation analysis of the effect of CK2 and GSK3β inhibitors, DMAT and SB-216763 respectively, on AR42-induced association of topoIIα with CK2α and GSK3β. Co-treatment with DMAT abrogated the ability of AR42 to facilitate the complex formation (Fig. 7E). In contrast, although SB-216763 blocked the association of topoIIα with GSK3β, it exhibited only a modest suppressive effect on topoIIα-CK2α interactions.

In vivo mechanistic validation

To confirm our in vitro findings of a functional role for the CK2α-Csn5-Fbw7 signaling axis in mediating HDAC inhibitor-induced topoIIα degradation, we conducted an in vivo study in a xenograft model. PLC5 tumor-bearing mice were treated for 3 or 6 days with a tumor suppressive dose of AR42 (25 mg/kg daily)(6). AR42 downregulated topoIIα and increased CK2α expression levels in xenograft tumors, without changing those of Csn5 or Fbw7 (Fig. 8A, input). Moreover, co-immunoprecipitation analysis revealed that AR42 enhanced the intratumoral association of topoIIα with CK2α, Csn5, and Fbw7, reminiscent of that observed in vitro.

Fig. 8.

(A) In vivo effect of AR42 (25 mg/kg daily; oral) on the expression and association of CK2α, Csn5, and Fbw7 with topoIIα in PLC5 xenograft tumors as determined by Western blotting and co-immunoprecipitation analysis. (B) Schematic diagram depicting the mechanism by which HDAC inhibitors facilitate the proteasomal degradation of topoIIα.

Discussion

In the literature, a number of stress conditions have been reported to induce the proteasomal degradation of topoIIα, including G1 arrest (34), glucose starvation (35), hypoxia (35), and adenovirus E1A-induced apoptosis (36), although the underlying mechanism remains unclear. Here, we report a novel mechanism by which HDAC inhibitors stimulate the selective degradation of topoIIα in HCC cells. As shRNA-mediated knockdown of HDAC1, but not other HDAC isozymes examined, could mimic the suppressive effect of AR42 and MS-275 on topoIIα expression, this drug-induced topoIIα degradation was, at least in part, attributable to the inhibition of HDAC1. Although HDAC1 has been reported to be associated with both the α and β isoforms of topoII (37), the significance of this binding in the effect of HDAC inhibitors on topoIIα degradation remains to be investigated.

We obtained evidence that transcriptional activation of CK2α expression represents a key driver for HDAC inhibitor-mediated topoIIα proteolysis. For example, ectopic expression of CK2α led to topoIIα repression, while pharmacological inhibition of CK2 kinase activity or shRNA-mediated silencing of CK2α expression protected cells from the suppressive effect of HDAC inhibitor on topoIIα expression. CK2 is known to bind and phosphorylate topoIIα on several serine and threonine residues near the nuclear export or localization signal (19,20,24). It was reported that CK2 could stabilize topoIIα against thermal inactivation in a phosphorylation-independent manner (38). Thus, this study provides a new insight into the role of CK2 in regulating the function/stability of topoIIα. Our data suggest that CK2-mediated phosphorylation of topoIIα, followed by GSK3β phosphorylation, facilitated its inclusion in the formation of a multi-protein complex with Csn5 and the Fbw7 E3 ligase, leading to its ubiquitin-dependent degradation (Fig. 8B). For instance, the silencing of either binding partner abolished the ability of HDAC inhibitors to deplete topoIIα, and pharmacological inhibition of CK2 kinase activity blocked both the formation of this complex and the drug-induced reductions of topoIIα levels. It is well documented that the Csn complex functions as a master docking platform to bring together a target substrate with its specific kinase and E3 ubiquitin ligase, which, in conjunction with the proteasome, facilitates the ubiquitin-dependent degradation (26, 29). The functional role of Csn5 in mediating CK2-facilitated topoIIα degradation is further corroborated by the reports that CK2 regulates the activity of Csn in mediating ubiquitin-dependent protein degradation (28), and that Csn5 is involved in topoIIα degradation in response to glucose starvation (27).

Fbw7, the substrate recognition component of the SCF complex, is recognized as a tumor suppressor because of its ability to target a number of dominant oncogenes (32). In this study, we used co-immunoprecipitation and shRNA-mediated knockdown of Fbw7 to demonstrate the functional role of Fbw7 as an E3 ligase targeting topoIIα. These findings reveal an additional layer of complexity in the regulation of topoIIα degradation and/or activity. Other E3 ligases have also been implicated in the degradation of topoIIα. It has been reported that Bmi1 is involved in topoIIα degradation in response to glucose starvation or the topoII trapping agent teniposide (VM-26) (31). In the present report, the role of Bmi1 in HDAC inhibitor-induced topoIIα degradation, however, was refuted by its decreased expression and lack of association with topoIIα in response to AR42 treatment (Fig. 6A). In other studies, Mdm2 (39) and BRCA1 (40) have been implicated in the ubiquitination of topoIIα, the former in the context of etoposide-mediated topoII degradation and the latter in the context of its participation in DNA decatenation. In addition, teniposide caused conjugation of small ubiquitin-related modifier-1 to topoIIα in HeLa cells, although its role in regulating topoIIα stability remains to be defined (41). The involvement of these pathways in HDAC inhibitor-induced topoIIα degradation remains to be investigated.

This study also reported the novel finding that topoIIα is a target of GSK3β phosphorylation, presumably at Ser1361, which might be primed by CK2-mediated phosphorylation at Ser1365, consistent with the reported mechanism underlying Fbw7-targeted protein degradation (32). Our data suggest that this double phosphorylation facilitated the recruitment of Fbw7 to the recognition motif ¹³⁶¹pSPKLpS¹³⁶⁵ at the C-terminus of topoIIα, leading to its ubiquitin-dependent degradation.

In conclusion, our report shows a novel pathway by which HDAC inhibitors facilitate the selective degradation of topoIIα, which underlies the complexity of the functional role of HDAC in regulating tumorigenesis and aggressive phenotype in HCC cells. Previously, we demonstrated the efficacy of oral AR42 in the in vitro and in vivo models of HCC through the inhibition of HDAC and modulation of multiple aspects of cancer cell survival signaling (6), which, as we now have shown, includes topoIIα degradation. As AR42 has entered Phase I clinical trials, the present finding may be of translational value for the use of AR42 as a component of therapeutic strategies for advanced HCC, in which systemic therapies have largely been unsuccessful.

List of Abbreviations

HCC

hepatocellular carcinoma

HDAC

histone deacetylase

topoIIα

topoisomerase II alpha

topoIIβ

topoisomerase II beta

DMAT

2-dimethylamino-4,5,6,7-tetrabromo-1H-benzimidazole

CK2

casein kinase 2

Csn5

COP9 signalosome subunit 5

shRNA

short hairpin RNA

siRNA

small interfering RNA

GSK3β

glycogen synthase kinase 3 beta

ChIP

chromatin immunoprecipitation

RT-PCR

reverse-transcription polymerase chain reaction